Science method

Optogenetics - Science method

Optogenetics is an emerging field of study that seeks to utilize photoreceptor proteins to manipulate neuronal cells through light stimulation. The practical applications could be far reaching including aspects of specific drug targeting in disease, direct gene targeting, and integrated fiber optics.
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We are planning to set up in vivo-freely moving optogenetic lab. We are using mice as animal model.
Most probably, we will order our staff from Thorlabs. However, I have a few small questions !
1) There are two sizes of ferrule and sleeve (1.25mm and 2.5mm) Which size ferrule and sleeve should I chose to make experiments in mice ?
2) There are also two size core diameter of optic fibers ( 200 and 400 mikrometers) Which size of optic fiber could be more eligible for mice situmalation ?
Thanks in advance
Best regards
Sertan
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Hi Sertan,
In mice the 1.25 mm ferrule is great. The 2.5 mm ones are really big, and you don't gain anything for using them in mice. Even with the 1.25 mm ferrules, it is difficult to do bilateral stimulation without angling the fibres - you need much more than 1.25 mm separation. If you're not sure how to plan angled fibres, I have developed an online tool to calculate angled stereotaxic coordinates: https://nicneuro.net/calculate-cannula-angle/
As for core size, I would also recommend sticking to 200 um. As others have stated, increasing the diameter will increase the absolute amount of light coming through the fibre. However, it will also disperse the light across a much wider area, so it won't necessarily help the irradiance, which is the critical value for activating opsins.
Finally, always use 0.22 NA fibres (even with LED's), as the higher NA fibres scatter the light too wide, with only a small increase in light output. Overall, your effective stimulation distance is reduced, check out my Optogenetics Depth Calculator for more info: https://nicneuro.net/optogenetics-depth-calculator/
Actually, one more thing: I would advise getting stainless steel ferrules. I found the ceramic ones seize up with the ceramic sleeves, and I ended up ripping some fibres out accidentally. Nowadays I just get these fibres from Thorlabs in boxes of 20 and cut them down to size myself: https://www.thorlabs.com/thorproduct.cfm?partnumber=CFML22U-20
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Dear RG-community,
in our lab, we currently use silica fibers (1 mm core, NA=0.22, multimode, 1 m) coupled to LEDs (470 or 630 nm) for in vivo optogenetics in behaving head-fixed mice. In search of fibers that increase optical power transmission, I have come across high NA plastic/polymer fibers (NA=0.68), which should obviously increase light power.
Besides this advantage for plastic fibers, are there any disadvantages to using plastic fibers instead of silica ones for our application?
Many thanks in advance!
Image source:
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Hi Yannik,
There's a lot to unpack here. Just to clarify, are you talking about the fibre optic patch cables (connecting the LED to the mouse) or are you talking about the optic cannula (that's going in the mouse's brain)? From your screenshot, I'm assuming you mean the patch cords. Doric do have some further info summarising the different types of fibre cores: https://neuro.doriclenses.com/collections/fiber-optic-cannulas/products/mono-fiber-optic-cannulas?productoption%5BFiber-optic%20Material%5D=Silica%20%2F%20Silica&productoption%5BReceptacle%5D=ZF1.25&productoption%5BFiber%20Tip%5D=FLT
Firstly, what optic cannula are you using? For optogenetics, I always recommend 200 um core 0.22 NA fibres, even when using LED sources. Reason being, as soon as you increase the NA of you fibre, you can get a marginal increase in light entering the mouse's brain, but it scatters far more widely, so it's actually less effective. For further info, check out my Optogenetics Power Calculator: https://nicneuro.net/power-calculator/
Next, about the patch cords. As Norman said, the optimal patch cord will depend on your LED. However, assuming that you are using a 200 um optic cannula, you do not need the full 1 mm diameter core. So, my suggestion would to drop the diameter of your patch fibre (maybe to 400 um), to maximise irradiance capture from the LED. In this case, the NA of the fibre shouldn't matter too much, so long as you do use the 0.22 NA fibre cannula.
Lastly, does your patch cable have a rotary joint? If the mouse is head-fixed that isn't necessary, and will lead to an avoidable loss of light power.
Unfortunately, many of the LED's sold for in vivo optogenetics are simply not bright enough. In particular, I'm not sure I've seen any outside of the blue 450-470 nm range that are really bright enough for normal optogenetic tools.
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Hi, I am planning to buy a stimulator, but thought the function generator - with the pulse generator option could be used instead as a cheaper alternative for triggering an optogenetic setup from Thorlabs. A function generator could also generate a train of pulses/ defined duration etc... Please let me know if we can do that and the pros and cons of such a setup. Thank you in advance.
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Thank you, Nicolas, I think Pulsepal will help my experiments.
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Hello there,
I am trying to assemble fibre optic implants for an optogenetics behaviour experiment, and cannot get the epoxy resin I am using (to fix the fibre optic in the ferrules) to set/stay set. I am using ThorLabs F112 Epoxy for Fiber Connectors, Long Pot Life (Eccobond F112 BIPAX). It comes as two solutions that you are expected to mix all in one go. However, as we only need a small amount at a time and I don't want to waste it, I have been trying to mix the solutions as 3 parts base, 1 part catalyst. Currently when I heat-gun the implants after assembling and let them sit over night at RT the epoxy sets as a firm gel that becomes more porous and liquid as days pass. Does anyone have any experience with this epoxy or other fiber optic epoxy? Am I using too much/not enough catalyst? Thank you very much for your input.
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I had something similar when trying to make fibre optic cannulae - the epoxy was just not setting properly. Either that or they were to viscous to get the epoxy into the cannula.
My solution was to stop trying to make them myself. Now I just buy them in pack of 20 from Thorlabs, and they cost less than £20 each. These are the ones I get:
Is there a particular reason you need to make them yourself?
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GtACR1 can be activated by light around 500nm to 600nm roughly and peak sensitivity is ~515nm. Activation of GtACR1 in neurons can inhibit neuronal activity.
I aim to activate GtACR1 for several hours to inhibit neurons expressing them.
Is it OK to constantly apply light to GtACR1 or I have to use pule stimulation?
I found several papers apply constant light to acitvate GtACR1 for up to 10 mins.
For over hours activation, I found on paper
(Mechanosensory input during circuit formation shapes Drosophila motor behavior through patterned spontaneous network activity)
It uses pulse stimulation 600ms duration with 1s interval. However, the author did not mention why they choose it (as least I did not see the clear explanation).
I know constantly activated Arch channels can become inhibitory and NpHR can become desensitized. So far I did no see any report talked about that GtACR1 can be strongly desensitized with constant light stimulation.
Any one has any idea should plus stimulation be used to activate GtACR1 for hours and what frequency should be used?
Best
Geng
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A new opsin has been developed that should perform exactly as you want. "Lamplight" is similar to the SSFO's that have a very long open time, which means that a single light pulse produces prolonged activation. Lamplight is, to my knowledge, the only inhibitory opsin that works in this way.
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I'm looking for the best strategy to obtain in vitro cells that once transplanted can be optogenetically activated and opto- or chemogenetically inhibited. However it is necessary that the same cell that is activated could be aslo inhibited, so I need a virus that contains both Chr2 and the inhibitory protein sequences. I only found the AAV eNPAC 2.0. It incorporates Chr2 and NpHR genes. Anyone of you has some experience with it? Are there any better?
For my porpouse the best inhibitory strategy would be chemogenetic, but I didn't find any virus that contains both chr2 and hM4Di sequences.
Any suggestions are really appreciated.
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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/
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I've made good optogenetic fiber implants for lasers in the past using lower numerical aperture (NA) silica core fiber from Thorlabs.  Recently, we've been working with LEDs which require a higher NA fiber in the 0.6 - 0.7 range. The highest NA fiber that Thorlabs carries is 0.5. Prizmatics has the right silica core fiber with high NA, but they only have core diameters of 200um & 250um.
Does anyone know a good vendor for this kind of fiber ( bare fiber, silica core, ~0.65NA)?
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I'm going to buck the trend here and suggest you do not use such high NA fibres, regardless of your light source. The problem is that any increase in NA makes the light coming out the end scatter more, decreasing your effective stimulation distance. With a higher NA fibre you need ridiculously high light output to maintain your effective stimulation depth.
For example, in a typical experiment with a 200um fibre and requiring an irradiance of 1mW/mm^2 to activate ChR2, if you wanted to have effective stimulation for 1 mm from the end of your fibre, you would need approximately 4.6 mW from a 0.22 NA fibre. But if you used a 0.65 NA fibre, you would need 27.4 mW. I ran these calculations based on the Aravanis model, and using my online optogenetics power calculator at: https://nicneuro.net/power-calculator/
My experience, using both Plexon and Prizmatix LED's, is that increasing the NA of your fibre only gives a small increase in light power. For example, I recently tested 0.22 NA and 0.50 NA fibres with a Plexbright blue LED and achieved 7.4 and 11.0 mW, respectively. In this case, increasing the NA produced a 50 % increase in light output, but due to the massive increase in light divergence gave an approximate 30 % DECREASE in effective stimulation distance in the mouse brain.
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Dear community,
before I lose too much time working with suboptimal software: which package is your current favorite for light-spread modelling in brain tissue?
Thanks for your help!
Michael
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Unfortunately, any system you use to model light propagation in the brain will be an estimate based on experimentally derived scattering values. And the scattering values are wildly different depending on who's done them (anything from around 9 to 40 mm-1).
A Monte Carlo model seems to be the best, but there aren't any simulations I can find that are easy to get quantifiable numbers from (as you seem to have also discovered - I have also used OptogenSim and it provides at best a very rough indication of light spread).
I have developed an online tool (https://nicneuro.net/optogenetics-depth-calculator/) which uses the Aravanis model. You select low, medium or high scattering tissues (depending on the relative amount of grey or white matter), input the parameters of your optogenetics system, and the calculator will give you an estimate of the depth you can expect to activate your opsin.
The important thing to keep in mind is that any light modelling tool will be an estimate, so I would advise using them as a guide to help decide your fibre placement relative to your desired site of activation.
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What would be a solid test to verify whether or not optogenetic stimulation of terminals causes antidromic stimulation of the soma and therefore activation propagating down other collaterals of that neuronal population? Ideally in an in vivo preparation...
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A possible in vivo solution would be to activate your neurone terminals (I like to do a solid 20-min stimulation), and then perfuse your mice 90 mins later and do immunohistochemistry for c-fos in the ChR2-expressing cell bodies. If you compare stimulated vs unstimulated, you should get at least an idea of whether you have activation of those cell bodies. It's not perfect, but it's quite easy to do and well-accepted in the literature.
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I‘d be interested in the quantification methodologies/pipelines that are used by different labs in order to reliably quantify the number of opsin-transfected cells in an injected ROI in case of somatic optogenetic stimulation in order to correlate w/laser powers.
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I usually count all cells in the expressing region and calculate the average cell density over the expressing region. Then you can use some calculator like http://web.stanford.edu/group/dlab/cgi-bin/graph/chart.php
to calculate effective power density in the tissue. If you know the electrical properties of your cells of interest you can approximate the power necessary for efficient modulation.
I can send you a chapter we wrote about how we do it. DOI:10.1007/978-1-4939-7417-7_8
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Hello
After transfecting my HEK293 cells with a construct containing an opsin and fluorescent reporter protein (e.g. ChR2-mCherry), I tried to patch one of my transfected cells based on red fluorescence of mCherry for electrophysiology and optogenetic experiments. Then, after patching and stimulating light, I notice that there is no current from this cell even though it produced red fluorescence under the microscope before patching
Have you experienced this kind of phenomenon?
Thanks.
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I have not experienced this in my work
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Hello
I am new to the field and I would like to ask on what is the criteria to say whether a photoswitchable compound or optogenetic molecule has fast kinetics and high spatiotemporal resolution at the cell-free model, cellular/in vitro model, in vivo and ex vivo model? Is there a consensus criterion to quantitatively qualify if a compound has a fast kinetic and high spatiotemporal resolution in these models?
For instance, if a compound becomes fully activated when turned on by light in less than 30 min, does it have fast kinetics?
On the other hand, if a compound can precisely activate certain neuronal regions in the brain but it has off-target activations in the surrounding regions around 20 uM from the region of activation, does it have high spatiotemporal resolution?
I may have mixed-up some terms here, I will be glad if this will be clarified in the discussion.
Thanks.
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That's the least I could do.
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Hello.
I'm currently studying the effect of optogenetic stimulation on social behaviors using the mouse model of autism spectrum disorder (ASD). In one of my experiments, I conducted the three-chamber test in two sessions, before and after the optogenetic stimulation. The social memory (or social novelty preference) test, which is the second phase of the three-chamber test, consisted of a novel mouse on one side and a familiar mouse that has been exposed to the subject earlier on the other side, as established by previous protocols.
In the first session (before the optogenetic stimulation), the subject mice couldn't distinguish between the familiar and novel mouse, which was demonstrated by similar time spent in interaction with each mouse. However, in the second session (after the optogenetic stimulation), the subject mice spent significantly more time interacting with the familiar mouse rather than the novel mouse (The target mice pair was replaced by a new pair in the second session). This result was quite unexpected and confusing since it is well known that mice generally seek novelty by natural instinct, which is exhibited by exploration time. But anyway, I still interpreted this finding as a positive improvement of the social memory, because after the stimulation, the subjects can now at least distinguish between familiar and novel targets.
Currently, I'm trying to search for some references that reported similar results of social attraction towards the familiar mouse, or any other alternative explanation based on the natural behaviors of mice, but haven't found any. Can anyone provide suggestions or recommend references with relevant findings? Is my interpretation correct?
Thanks in advance.
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Thank you for your answers.
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Hi, I am Joe and currently, I am working in a neuroscience lab using the optogenetic technique. And I have some technical problem with that, which the AAV always injected off target, for example when I inject the AAV in the BLA. However, there is always a lack to the CeA and the injected passage which bothers me a lot.
Therefore I would like to ask if there are any tips to prevent the situation that I mentioned? Thank you so much for your help.
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As suggested above, you should try lower volumes and injection speeds. Also, you could aim to the lateral portion of the BLA if leakage into the CeA is your main concern.
Another thing to consider is how you define a "leak"? If you are simply looking at fluorescence intensity with a relatively small magnification then you might mistake labeled axons coming from the BLA as a leak. This is not so trivial to solve with some vectors, if the fluorophore is directly attached to the opsin (and therefore does not fill cells, but rather outlines their membrane). Nevertheless, as a first step, I would suggest staining for NeuN and then using either a confocal or high magnification in a fluorescence microscope (e.g., 20x) to see if it is in fact the CeA somata that are expressing your opsin, or simply a high concentration of opsin-expressing axons from the BLA.
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Currently ,I am working in a neuroscience lab.And I am going to test the neurotransmitter level under the optogenetics activation but also observe the calcium signal for mice
So I would like to ask if it is possible that I can use those techniques at the same time for the real time manipulation and observation? thank you so much
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Dear Joe, as an inorganic chemist I'm clearly not an expert enough to give you a qualifed answer to your interesting technical question. All I can do right now is suggest to you the following potentially useful article which might help you in your analysis:
Tonic or Phasic Stimulation of Dopaminergic Projections to Prefrontal Cortex Causes Mice to Maintain or Deviate from Previously Learned Behavioral Strategies
This paper is freely available as public full text on RG. Please check particularly the "Materials and Methods" section on page 2.
Good luck with your research!
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Hello everyone!
We are having this issue when transfecting rat cortical neurons using lipofectamine and niosomes. We are transfecting the optogenetic plasmid CAG - ChrimsonR - TdTomato, and others like Catch - YFP and even GFP.
We are having this problem in all of them.
It seems an apoptotic/necrotic reaction, and we'd like to know if anyone has experienced the same problem and how to solve it.
Thank you!
Kind regards!
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I have an update for you guys.
We are using ChRR - Syn now. I thought it could be something about the promoter (CAG) but it doesnt seem so. It's happening again
The pic is a neuron transfected with lipofectamine 2000 and ChrimsonR-Syn at 2:1 ratio and it's clearly dying. However it seems that the cells last a little longer in a "healthy" state with ChRR-Syn.
Best Regards!
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In fiber photometry experiments we see a decrease in our 405 nm signal during increases in GCaMP (465 nm) signal. Additionally, we have been trying to do photometry simultaneously with optogenetic stimulation in a nucleus which sends glutamatergic projections to the nucleus in which we are doing fiber photometry recording. When we deliver stimulation, we notice that there is a decrease in our 405 nm signal. The fluorescent reporter we are using in these experiments is not a calcium indicator however. Does anyone know what causes these drops in 405 nm signal? Could it have something to do with pH changes during neuronal firing? Is it some sort of interference between channels? Any help is much appreciated.
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Hi Andrea,
We haven't actually tested different intensities for the optogenetics stimulation. My understanding of photobleaching however is that it is a continuous decay in the signal. In these experiments we deliver optogenetic stimulation for 24 seconds and then it turns off. The decrease occurs during those 24 seconds, then the signal rises back up.
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My in-vitro culture consists of E18 rat cortical neurons. I am looking to quantify activity with calcium using GECI, RCaMP1a. The optogenetic experiment relies on the excitation of ChR2 transduced cells. Therefore I have to dual transduce my culture with both RCaMP1a and ChR2. The cell density in the plate is 150,000cm^2 in 1mL of media. The ChR2 is AAV9 and the RCaMP1a is AAV1. I usually transduce them by choosing the titer 1E10 for RCaMP and 2E10 for ChR2, and then adding them to 25uL of media and pipetting the solution directly into the wells. What I am finding is that the dual transduction wells are not as active, if at all, as the single transduction RCaMP wells. Has anyone encountered dual transduction affecting cell health and activity? If so what titer do you suggest using and is there a better way to dual transduce my cultures that would prevent this from happening? Thanks!!
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I've experienced the same thing, no activity all. But the neurons appear healthy and in perfect morphological state. You are not alone... I've also co infected with Gcam and Chrimson, and the same thing....
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My lab uses the Plexon optogenetic system, but we had an issue during our first experiment with the patch cable twisting, then breaking, resulting in a huge loss of light output. In addition, our mice do not seem to generate enough force to rotate the mounted commutator. Has anyone else had this issue/does anyone have a picture of their setup? Any help is appreciated.
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I hope I'm not too late to help you with your issue. We also use the Plexon optogenetic system, and have encountered the same issues as you. The good thing about the Plexon system is the fewer optic connections which results in less light loss and a strong light power out the end of the cable. As you say, the commutators are problematic and their optic cables are very fragile, though overall we've found it to be ok. Here are some things we do:
  • Bilateral optic fibre connections are better than unilateral, as the torque on the commutator can cause the single fibres to twist and shear.
  • We use 0.5m cables, with the height of the mouse cages adjusted to provide only a small slack. This prevents the cables from over-twisting, and encourages the commutators to spin when the mice turn. It is still worthwhile having someone present to spin the commutators if needed to relieve torque, although we find this is rarely needed.
  • Plexon have recognised the problem with the heavy commutators and are now selling more durable versions of the optic cables (we have yet to get any, but they are on our buy list).
Unfortunately, there are limited options for in vivo optogenetics without the use of lasers (which we wanted to avoid for safety reasons). Out of the LED systems, I think Plexon does make a decent compromise between usability and light power outout, whereas I'm not sure what the actual light output of the other LED systems are, as the manufacturors are reticent to publish that information
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Hi all.
Recently, I encountered a problem regarding the optogenetic stimulation experiments.
In our study, we are using the blue laser (473 nm) to optogenetically stimulate parvalbumin (PV) neurons at 40 Hz (gamma frequency) in the mouse basal forebrain. According to the principles of optogenetics, the neurons that do not express light-sensitive opsins (ex. ChR2) are not supposed to be responsive to optogenetic light delivery. However, we discovered that even in the control virus (GFP)-injected group and the Sham (no virus injection) group, the 40 Hz response was observed when the blue laser was delivered at 40 Hz, just as in the ChR2-injected group (although this response was weaker compared to the ChR2-injected group). Other stimulation conditions were the same in all groups. (The laser intensity was 70mW/mm^2 in all trials.)
We hypothesized that this can be due to heat-inducing effect of optogenetic light delivery into neural tissues according to our literature survey, since the blue light is also known to induce heating of nearby tissues, which can actually induce changes in neuronal activities. However, we also questioned that if this non-specific stimulation was due to heat induction, how can the temperature increase during the stimulation interval induce neuronal firing at 40 Hz just as in optogenetic stimulation, rather than non-specific firing at random frequencies (i.e. Can the temperature change of neurons take place every 25 ms?).
Has anyone ever faced the similar issues?
I wonder if there could be other possible explanations or causes for this phenomenon.
I would also like to inquire if there are any references or resources that may deal with this problem.
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@Justin Siemian ,
Thank you for your advice! I will check that out as well.
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If a neuron expressing an optogenetic actuator is light-stimulated in the soma, will this lead to neurotransmitter release of this same neuron?
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Hi Soledad,
depends on the optogenetic actuator. If you use an excitatory one this will lead to the release of the neurotransmitter of the affected cell. However if you use an inhibitory one (e.g. Halorhodopsin or Archaerhodopsin ) you hyperpolarize the cell and therefore supress its firing and transmitter release.
I hope this is of some help.
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Can anyone made me clear the way of photostimulation under 2P optogenetics spiral scanning?
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Follow
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We are looking to inject optogenetic contructs with ArchT or eNpHR3.0 in ACC and we are considering several options.
We are curious what is your experience with the different AAV serotypes, promotors and volumes of injection in the rat cortex and your most trusted suppliers
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I would look around the new addgenen database
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Hi, I'm recently trying to validate c-fos antibody staining with optogenetics stimulation.
The wild type mice was injected Syn-ChrimsonR-tdtomato virus with cranial surgery.
(somato-sensory cortex, 5mm diameter)
After curing period (3-5weeks) I checked the virus expression by 2P microscopy.
Almost mice were confirmed that they expressed tdTomato and I gave the stimulus light in to the mice cranial window site.
Before giving the stimulus light, the mice were awake and I put them into the isoflurane chamber and anesthetized. After that, I delivery the stimulus light.
(674nm laser was used. 50mW at the end of tip in continuous wave, 10Hz stimulation duty-cycle 10%, 1min stimulus and 1 min rest...X5)
After stimulation, mice were placed into the cage and rest at least 75min (Waiting c-fos expression time).
After rest time, the mice were operated perfusion.
The next,
1- 1day PFA fixation
2 - 100um slice by vibratome
3 - 2h Normal goat serum blocking solution embedding
4 - Primary antibody solution embedding ( Merk anti c-fos Rb) (3day)
5 - Washing by PBST 30min/3th
5 - Secondary antibody solution embedding (Alexa 488 - anti Rb)
6 - Washing by PBST 30min/3th.
But all the IHC imaging, the c-fos was formed filament structure in L1-5 in gray mater and nearly existed in soma.
Furthermore, the c-fos signal was very well distributed in whole brain region (gray mater, white mater...wherever!!)
The distribution phenomenon was same If I added another antibody protein (NF, anti-tdtomato,..)
As I know, the c-fos could not escape the nucleus.
Why the c-fos signal was existed in every where?
(I never see the filament like c-fos signal)
And why the c-fos signal was well distributed whole brain?
+1) I checked the cross talk(bleed through) of fluorescence and there was little signal cross talk that could be ignored.
+2) The mice was not trained for the light stimulation . - I think this could be an clue for this problem but I have no confidence.
In my opinion, even they had a surgery event, grabbing would affect mice physiology (noble experience).
i.e : dread, fear, stress -> it could be classified as emotional stimulation and run or resist would be induce motor and somatosensory stimulation.
I also attached the image for that. (blue : NF, Red : c-fos)
Thanks,
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Florian Hetsch
Hi there!
Recently I solved the trouble!
As you mentioned before, the filament structure was dendrite!
I changed the stimulus source as 4-ap (potassium blocker) induced seizure model.
And finally found that the previous optogenetic stimulus was not enough to induce action potential!
In 4ap model, ipsilateral site had less c-fos positive cell body (may be spontaneous firing) shape but also shows the dendrite structure.
On the other hands, the contralateral site showed a lot of c-fos positive cell body shape near the 4-ap injection site and dendrite structure.
In my opinion, the c-fos antibody which I used was low affinity or weak secondary binding thus I observed the dendrite.
However when the certain stimulation was delivered, the c-fos antibody was enough to distinguish between stimulated and non stimulated neuron.
Now I would change optogenetics stimulation parameter!
Thanks a lot !
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Please provide solution that is readily available. Also what is the downsides of each of them when comparing to electrophysiology?
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you will need to ensure that the excitation spectra of both the indicator (voltage or calcium) and the opsin are sufficiently separate so that you don't have any crosstalk, i.e. the imaging wavelength exciting the opsin.
Our lab found a solution using Chronos for the opsin and CaSiR-1 for the calcium indicator
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When you cut a slice, you sever many axons. So when you use your stimulating electrode (or a light in optogenetics) to evoke currents in the cell you've patched, can you evoke a response if the afferents are severed proximal to your stimulation?
What I'm hung up about really is, if you express channelrhodopsin in some nucleus, then cut a brain slice which does NOT include the cell bodies of that structure but does have the afferents, will the light evoke a response from those severed axons?
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Most definitely. For example, we and others express ChR2 (or derivative like chronic) in prefrontal cortex and after letting the neurons express for 4-6 weeks, make slices and can optically evoke TTX sensitive monosynaptic EPSCs in brainstem targets (e.g. dorsal raphe). These EPSCs can also be reinstated following TTX blockage by adding 4-AP which, by blocking K channels in the cut axons, allows the Opto depolarization to be large enough to open voltage-gated Ca2_ channels and drive synaptic release. Thus, the cut axons as sufficiently hyperpolarized to sustain Na-channel dependent action potentials.
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Hi,
I would like to begin to learn optogenetics in a systematic way.
Because I have the opportunity to work with a femtosecond laser system, I am mainly interested in the optical part of this issue. Do you have any suggestions on where to start? All suggestions are welcome.
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I think the best way for you to acquire knowledge and skill to understand optogenetics studies and to plan yours is to read recently published papers. First, you may need to focus your study on a specific function or dysfunction of the nervous system, and where in the brain you would like to see the effects of your experiments.
Then you can narrow down the papers that would help you to implement the right protocol for your stimulation and recordings.
Try the following to get a clue:
A pioneering paper on optogenetics by Prof. Deisseroth:
another one:
Development and application of optogenetics by Prof. Deisseroth team:
There are also occasional webinars related to optogenetics on the web.
Search for possible online courses too.
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I need an AAV to introduce channelrhedopsin into glutamatergic neuron. This should be able anterograde tracing all the way to the endings so that the endings can be activated by photostimulation after being sliced.
Thank you!
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This vector from UPenn works well
AAV1.CaMKIIa.hChR2 (H134R)-mCherry.WPRE.hGH
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Hi.
I've been using LED for optogenetics and calcium imaging.
For now with the devices, I'm only able to adjust the currents or % power of the LED as out power can vary from patch cord to patch cord.
While it seems like people usually adjust the power of LED by W/mm^2, how would I be able to measure that?
What kind of meter do I need?
Please recommend me if you've been using one.
Thank you!
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Dear Kwanghoon,
I also can recommend the power meters and sensors recommended by Roni Hogri and Niraj S. Desai above (I find the S130C slim sensor quite useful).
However, these devices measure *power*, not intensity. In order to measure intensity with these, you need to know the total illuminated area on the sensor. That you can achieve in a number of ways, depending on what type of light source you are using.
1. If you have a uniform beam that has a larger diameter then the sensor aperture (9.5 mm diameter for the S130C), then you can estimate intensity as total power divided by sensor area (i.e. pi*r^2). Caveat: if you beam is not uniform, this estimate will be inaccurate.
2. If you have a uniform beam that is several mm in diameter, but difficult to measure the diameter exactly, you can place a smaller aperture onto the sensor, to limit how much light reaches the sensor. For example, if you drill a 3 mm diameter hole into piece of black material and place this over the sensor, and your beam is > 3 mm in diameter, then this would allow you to calculate the intensity as described in (1) above.
3. If you are illuminating a sample through a microscope objective, then these methods may not work well (especially if using a high magnification objective that typically generates a spot smaller than 1 mm). In that case, you would need to calculate (based on objective properties, wavelength, etc) the effective beam waist diameter at the focal plane. Use this together with the total power measured to obtain intensity.
4. If you are using a fiber to deliver the light, then all you need to know is the fiber core diameter (e.g. 0.2 mm) and the total power. As others have described, you just need to make sure that all of the light out of the fiber reaches the sensor, to get a measure of total power. Be careful because the fiber tip can scratch the sensor surface if they come in contact. In this case, you are estimating the intensity *at the tip* of the fiber. The intensity at the sample will most likely be much lower and generally quite difficult to estimate. But see:
That website is currently unavailable, so here's a snapshot from 6/2018:
For consistency, if you are doing in vivo experiments with an implantable fiber, I recommend measuring the total transmitted power *prior* to implanting the fiber "cannula", as this can vary from implant to implant (and hence animal to animal). You can then adjust the command power to achieve the same power delivered to the brain in each animal.
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Hi all,
I am starting a new projet in which I'd like to specifically modulate GABAergic transmission on acute hippocampal slices preparation by optogenetic stimulations.
I have no experience in such assays and I'd be greatful if some of you could give me some advices, specially regarding to the Equipment required.
I already have a functional extracellular recording set-up. From what I Believe - just talking about material Equipment no mice/slice themselves - I could " just" direct a laser to the slice recordings chamber and record "as usual".
In the litterature, I've found that Diode-pumped solid-state lasers (DPSS) are more suitable as light source, Can anyone confim?
Does anyone knows if combining the laser source with an optical fiber we can submerge the light source into aCSF for higher spatial accuracy ?
Any laser or other Equipment supplier recommandations are warmed welcomed...
By advance, thank you !
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Thank you both for your help. It will help a lot
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It´s normally atributed to photoelectrochemistry but most of the examples are with semiconductors, where the required photon energy is smaller than a metal. Also when a electrode potential is applied it will decrease the required energy for the generation of photocurrents.
My setup doesn´t have any additional electrode potential. My electrodes are submerged in electrolyte (100mM NaCl) and are illuminated perpendicularly to the surface (625nm @ 6mW/mm^2).
I´m not using semiconductor electrodes so there shouldn´t be current in non UV wavelengths. I´m using different MEA (Au, TiN) but I´m still observing this current once the electrodes are illluminated, also if I illuminate just the tracks I see a voltage.
In the following article[ ] they explain the required energy if the difference between the Work function of the metal and the solvation energy of the electrons. The problem is that they don´t give any proof for this hypothesis.
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Hi David,
It's the photovoltaic effect, or the Becquerel effect (not the photoelectric effect). The photoelectric effect depends upon applied electrode voltage, the photovoltaic/Becquerel effect does not. I'm told there can also be an impact of the photoconductive effect, where light makes a material more conductive, which can also creates a voltage difference.
Anyway, the point is, it the effect occurs across the visible spectrum and down into the near IR.
It is a very common problem in optogenetic experiments. If you do a search you'll see lots of papers discussing it (e.g. Kozaia and Vazquez., 2015) . Theoretically, there are certain coatings that can be applied to electrodes (e.g. ITO) that will prevent the effect, but I don't think many people go to that extent.
If you're using MEA, you're actually quite lucky, as your light induced artifacts should be very reproducible from day to day, hence you can generate a recording of them with no cells on the array to generate a pure artifact, and then simply subtract this signal from your data.
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Hello everyone:
I'm now trying to model some optogenetic experiments. But I'm struggling with propagation of the potential. The modeling requires relatively precise description of the spatiotemporal dynamics of electric potentials. Here is an example question: consider a spherical soma, if I inject current into one point of it, how potential of each patch in this spheres changes in time?
On the internet it's almost exclusively cable theory that provides underlying modeling. Although it is absolutely suitable for most of the events in neurons, it may not be enough precise for ultra-fast source of current that changes in space within micro-second level. I would say before the iso-potential status is reached the source of current has left so old integration process ceased and new integration process occurs in another place. So a more precise model is required here.
I'm not a mathematically or physically intelligent guy so maybe some considerations are wrong. But any suggestion will help! Thank you in advance!
Best
Yao
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> though it is absolutely suitable for most of the events in neurons, it may not be enough precise for ultra-fast source of current that changes in space within micro-second level
It is precise enough. Somata are isopotential on any biologically relevant timescape. You can model it yourself if you don't believe Rall and others. NEURON will happily split a soma into hundreds of compartments.
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I just started to do electrophys (patch-clamp) experiments from old (> 3 month) CD1 mice, but I'm struggling to get any recordings from subthalamic neurons.
Any suggestions on how to improve our brain slice prep, specifically for getting healthy subthalamic neurons?
  • Patching is not the problem because I can get good recordings from young mice and rats.
  • I've read the work by Jonathan Ting, but haven't found any information about the subthalamic nucleus specifically.
  • I currently use NMDG+NAC cutting (5 ºC) and recovery solution (12min @ 34 ºC)
  • Incubation and recording solutions are normal ACSF
  • Transcardial perfusion didn't seem to help
  • CompressTome didn't seem to help too much
Has anyone recorded from old subthalamic neurons?
Any help would be greatly appreciated :)
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Hi Alberto,
Thank you for your response.
We have started adding HEPES to the incubating solution. The results are not very positive at this stage. See attached file, for the image of the STN in a slice that was incubated with HEPES solution.
Hi Ted,
Thank you for your response.
The neurons in other regions look better than the STN neurons and can be recorded from, unlike the STN neurons. See attached file for comparison between the SNc (substantia nigra pars compacta) and STN. It appears that STN neurons are perhaps more vulnerable to damage.
Thanks for sharing those references, we will take a look at them.
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Hello to everyone,
my name is Stylianos and these days I have focused on pairing recordings.
I cause 5 APs (5 pulses 20Hz) to a SOM-Martinotti neuron and I record IPSCs in a principal neuron in cortical slices at 0 mV in presence of DNQX and CGP. I found depression of the synaptic transmission (pulses 2,3,4,5 were smaller in amplitude compared to the 1st). This finding is in agreement with the literature.
BUT.....When I did the same experiment by optogenetically activating many SOMs with CAG.ChR2 and recording IPSCs in principal neurons, I found facilitation of the 2nd pulse compared to the 1st (in these experiments I applied the PPR protocol with 2 pulses at 20Hz).
So, I am wondering if there is an explanation about this discrepancy.
I thank you a priori for reading this.
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Interesting question.
I don't KNOW the answer. But I have a testable hypothesis.
My theory is that with optogenetic stimulation you are basically seeing temporal integration in SOM cells. That is to say, with your first light flash, you are causing say, 20% of SOM cells to spike, and the rest to be depolarized. On your second light flash, you are causing 40% of cells to spike, the increased number caused by residual depolarization left over from the first optogenetic exposure (i.e. the membrane is still charged up).
If this is the case, you should notice a change in the paired pulse ratio with varying light intensities, i.e. with very weak light intensities, you should see more facilitation than with very high intensities. Furthermore, the facilitation should not behave to changes in extracellular Ca2+ concentration in the expected way (though the first test is easier to perform), i.e. if you increase the extracellular Ca2+ concentration you should see no significant change in the paired pulse ratio.
Interested to see if anyone else has a better idea.
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I have a fluorescent light source and an objective lens and would like to measure the light intensity at the focal plane. I can measure the light intensity at a different position but need to calculate it at the focal plane. I am using a 40x, 1.3 NA, 0.2 mm WD lens. I guess I need to make an area calculation.
For context, I am using light to stimulate cells (on a coverslip) expressing an optogenetic construct and need to be able to estimate light intensity.
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The first step is a light power meter, like a PM100D from Thorlabs with appropriate detector. Set that to the right wavelength, and read off the power in mW.
The you need to estimate the size of the excitation area. You probably have an etched calibration slide somewhere. Look down the scope, see what the diameter of your field of view is, and use 3.141*D to know your area.
Divide the first measurement by the area, and your done.
Yes, the measurement of your excitation area will not be 100% accurate, but it will be close enough.
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Hi, I'm looking to use the new Ai162 transgenic reporter to cross into my PV-Cre driver line for wide field calcium imaging (Ai93 line does not express well enough in interneurons). Is anyone using this mouse line yet or know where I might acquire it? Thanks! 
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Is there any new development on this issue in terms of mouse lines?
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Has anyone used collagenase to thin dura in NHP such that chronic, drivable electrodes can be utilized?  
If so, what type of product did you use?
How did you apply it?
What was the concentration?
How long did you allow it to sit?
Any help would be much appreciated!
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Hello, have you successfully used collagenase to weaken the dura in the NHP?
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Could you advice virus with eArchT3.0 light-activated pump which can be injected to wild rat/mice? We need expression of eArchT3.0 in neurons in entorhinal cortex, non-specific to the type of neuron.
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Thank you, Thiago! That is what we need. I found on the internet "Databinge AAV Presentation" of Matthieu Vanni and Alexa Nelson. And using their recommendations I also chose UNC (it seems to be cheaper) and viruses AAV-hSyn-eArch3.0-EYFP or AAV-CAG-ArchT-tdTomado. Maybe Synapsin promoter is better (that to exclude the expression of rhodopsin in glia) but I heard that EYFP is not always visible. Thank you for your help. We will try AAV-hSyn-eArch3.0-EYFP virus.
Best regards,
Lena
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Optogenetics is branch of biotechnology, which is growing and emerging in the treatment of many brain diseases including depression, Parkinson, so what are new innovations related to obesity and diabetes.
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Optogenetics is an emerging field that uses light-sensitive proteins to regulate biological activities in the body. The technique has been envisioned as a way to treat a range of diseases, including Parkinson’s and schizophrenia.
Optogenetics to precisely control cells to deliver insulin to diabetic patient. The approach could be used to continuously monitor blood glucose levels in human diabetics and automatically produce necessary insulin, a hormone that converts sugar from food into energy the body can use.
The researchers engineered human cells with a light-sensitive gene that is found in plants and produces insulin on cue when activated by wirelessly powered red LED lights. They inserted those lights and the designer cells onto small, flexible discs that were then grafted onto the backs of person or any place of the body.
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I am trying to figure out a way to have cortical slices with detectable only the SST Martinotti cells. And if possible a way (with virus) to optogenetically activate only these inhibitory projections to principal neurons. I would appreciate any help in this matter. Thank you.
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The GIN mouse line (JAX 003718; Oliva Schwann JNeurosci 2000) exclusively labels Martinotti cells (MCs) in layer 5 of visual cortex, see Buchanan et al Neuron (2012) 75:451. It is a very sparse line, GAD67-based so post-hoc characterised as MC specific, it only gives you the tag, not the ChR2, and the genetic background might not be the best (albino), so perhaps not ideal for your purposes. To my knowledge, there is no Cre line specific for MCs. Any Sst-Cre line will drive at last three cell types, see McGarry Yuste Front Neural Circuits 2010, doi: 10.3389/fncir.2010.00012. I suggest the GIN line + patching + post-hoc verification with morphology and spiking pattern to achieve the best specificity for MCs.
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I am doing an experiment that involves injecting virus expressing mCherry-tagged channelrhodopsin under the synapsin promoter (AAV-hSyn-hChR2(H134R)-mCherry) into a mouse containing a GFP reporter for my cells of interest (glial cells, so no neurons should be labeled). The intent is to excite neurons (which should be the only cells expressing ChR2/mCherry) and assess the effects on surrounding glial cells, which should be expressing GFP.
However, I ran into an issue where I get profuse labeling of neurons when I perform GFP antibody staining, specifically in areas that have been infected by the virus. I have confirmed that this seems to be some interaction between my GFP antibody and the ChR2 virus, as even wildtype mice injected with ChR2 virus that should have no GFP labeling have the same non-specific staining.
I have tried two different kinds of GFP primaries (rabbit and chicken) and both have given me the same result. I also thought it may be non-specific binding to the mCherry protein, but other Q&A topics on ResearchGate seem to suggest that mCherry and GFP are quite different proteins, and you wouldn't expect such an interaction. Finally, it is perplexing that the neurons labeled by the GFP antibody are usually not the same ones positive for mCherry.
Does anybody have an idea what might be going on here? Any help would be greatly appreciated.
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Suggested experiments for determining if problem source is: primaries, secondaries, GFP cell marker misdirected to wrong cells or, virus. Primary Ab omissions are always useful to rule out/in some problems.
1) GFP primaries on sections from untreated wild type mice, secondaries as usual (2 slides, one rabbit and one chicken)...do you get label?
2) Primary omission on sections from untreated wild type mice, secondaries as usual (2 slides, one rabbit and one chicken)...if label above is it just due to secondary?
3) GFP primaries on sections from untreated (no virus) GFP mice, secondaries as usual (2 slides, Rb and Chicken).....glia and neurons?
4) Primary omission on sections from untreated (no virus) GFP mice, secondaries as usual (2 slides, Rb and Chicken ).....if neurons label in 3, is it secondary?
5) and 6).....as above 3 and 4 but GFP animals with virus injections (4 more slides total)
Notes: 1)use all fresh buffers and blocking solutions. 2)If slides are dear, just do 1-6 using only one or the other (Rb or chicken) primary Abs with its secondary Ab.
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We would like a 2P optogenetics rig for as cheap as possible. Ti:Sapph are nice tunable lasers but not really cheap. Ideally we would want a femtosecond pulsed laser about 920nm, but lasers are a lot cheaper around 1040nm.
It looks like that the excitability of ChR2 is low but not zero at 1040nm. I am not sure if this means 1040nm is suitable for ChR2, or maybe it would require power levels that exceed safe levels for the animal. Has anyone tried this?
I know we could use 1040nm with a red-shifted opsin, but we would prefer not to.
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I am aware my comment may not be extremely useful as I am not familiar with the details of lasers used in 2P optogenetics, and not too familiar with the prices of Ti:Sa lasers.
However, I still want to ask: have you considered using a 1040nm laser to pump an optical parametric amplifier? Perhaps such a setup would still be within the budget? 920nm is a wavelength easily accessible for an OPA pumped with the SH of 1040nm lasers.
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I plan to use a 1W laser for research about optogenetics, is there any special requirement for patch cable connected to high power lasers? Thank you!
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If you are using a laser with near-infrared emission, you don't need to warry about the damage problem at the output end facet. However, at the input facet I suggest you use a fused splicing method, rather than direct  mechanical connecting or too tight a focusing. If you want to launch a free-space beam into a fiber patch cord, you should use a NA-adapted focusing component.
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Are there any labs that work with optogenetics in Turkey (Ankara, Istanbul)? So far I looked through the Pubmed and Google, but I couldn't find anything.
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Take a look at this lab: http://denizatasoylab.com/
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After transducing a culture of primary neurons, is there a quantitative way to determine how effective was the transfection? I will transfect the neurons with a dual EGFPRFP fluorophore, using an AAV2 serotype 
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 Hi Ariadne,
I think that counting fluorescent cells and dividing by fluorescent cells plus non- fluorescent cells is pretty standard. Neurons are pretty easy to pick it in culture, so unusually, people just pick a few fields of view to make the quantitation.
FACS is more accurate, but then you have to worry about glia. You might try NeuN staining to isolate the neurons, though, if you decide to go that route. 
Best of luck, 
Joey 
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The techniques I've heard of are:
1. Switch to glass electrodes (not compatible with my experiments)
2. Play around with relative positions of fiber optic and electrode (all locations I've tried still end up with artifact -- but we are very constrained in space so not much flexibility)
3. Record the light artifact alone elsewhere in the brain and subtract it out before doing spike sorting. 
Is #3 really reliable? Any issues to be aware of? Any other ideas/other metals/coatings that might help?
Details: I'm delivering short pulses of light (473 nm, 2 ms) in awake, head-fixed mice and recording the response in ChR2-expressing neurons. Currently using tungsten wires (1.5 MOhm) coated with parylene-c. Light is generated by a laser and delivered via a fiber optic, inserted through the same guide tube as the electrode, and positioned 0-2 mm back from the tip of the electrode. 
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 In order to find out whether Refik is right and it is an "electrical artifact", did you trigger the laser, but had unplugged the fiber before, so no light reaches the tissue? And I really mean UNPLUGGED aka physicallyY BLOCK the LIGHT?  If you still see an artifact, the method of Refik might work... or some more thoughts about ground connections in your setup. 
If it really is the light itself - is it intensity dependent? If you need to stick with single wires, go for stainless steel or Pt (FHC and many others...) or glass coated PtIr  multisites (AlphaOmega or Thomas Recording). We do see light artifacts when illuminating MEA arrays - but I doubt it is a very specific photoelectric effect...  More of a thermal-electric effect (thats why I was asking about the light intensity dependence...)  
If you go to apply silicon probes, be aware, that (very much dependent on the lithography process), the interface between metal and electrolyte might affect the semiconductor below. It may very well be a depletion layer down there, thus establishing a real photodiode perpendicular to the electrode into the Si-bulk.... Thats why I prefer Si-probes based on the Silicon-on-Insulator over p-doped Si-probes.... ;-)  So, I am not quite sure which process Neuronexus is using for all their probes (it is recognizable from their spec sheet), but you might want to look into the Atlas Si-Probes as alternative... Their newest ones should not show these photo-currents. 
And last but not least, refering to template-subtraction: An artifact is an artifact and if it is big enough to be a nuisance, it's subtraction is big enough to spoil your data. 
I know everybody is using this method, but I am not really fond of it - but alternatively blanking your signal is really worse!
Good luck!
uli
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I'm looking for the most reliable way to selectively activate axon terminals of dopamine neurons projecting from VTA/SN to specific areas of the striatum (subregions within NAc or caudate/putamen) in behaving mice.
Temporal & cell type (i.e., only DA neurons) specificity are crucial to my experiment, so optogenetics seems to be the way to go. Because it seems extremely tricky to get a robust enough (i.e., behaviorally relevant level) ChR2 expression that's limited to DAergic (DAT+ or TH+) AND projection-defined neurons, I'm trying to see if the approach of expressing ChR2 in VTA/SN DA neurons and then light-stimulating terminals in specific striatal regions would be more feasible.
I saw some studies in slice (e.g., https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3602228/ and https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4537642/) that delivered light stimulation to striatal DA neuron terminals to induce DA release, but I'm not sure how well this would translate to an in vivo setup. Has anyone got this or something similar to work? Are there any caveats I should be aware of?
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Hi 
I would suggest you to use DAT-cre mice since TH is not exclusively expressed in DA neurons. 
There have been several studies using transgenic mouse lines, injecting with viruses in VTA and then stimulate with optical fibres the terminal areas to study behavior. You can have a look on those for example:
Good luck!
Zisis
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There was a recent NIH videocast lecture discussing the ventral v dorsal model of the visual system. My impression is the model was debunked in line with more massively parrale and network models of brain functions.
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The ventral vs dorsal dichotomy originates from some differences in object processing correlating with the lesion and functional activation studies, e.g. prosopagnosia with right  temporobasal lesion / activation focus and pure alexia in the homologue regions in the left. Positional tasks and spatial tasks comprising tasks with respect to the own body are correlated to dorsal lesions / activations. From a clinical / lesional pov, already the cerebral akinetopsia (lesion at the lateral temporooccipital junction) raise some concern. In my opinion, connections from primary brain areas in a modality to other modalities should be more accounted for, and of course there must be a connection between dorsal and ventral pathways, which makes this dichotomy a good similar kick-off to test hypothesis' against as it was the case in language localisation.
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I'm trying to implement the use of rabies virus for tracing and optogenetics purposes in the university that I'm working. Does anyone have the approval to work with rabies in Netherlands that could give me the orientation in the bureaucracy process? 
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Have some idea from here too..
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Specifically, does long term red light exposure (~12-24 hours) influence post exposure vision? I'd like to do some visual attention assays following optogenetic activation, however, my controls (not provided all-trans-retinal) are performing abnormally in visual behavior assays, which suggest that the red light is having a secondary effect.  
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Hi Chelsie,
I have no experience investigating fly vision or long-term red light exposure, but I have used CsChrimson extensively over the past 2 years, both in explant brains (preferably) and in vivo, and might have some clues to what you're seeing.
First of all, my colleagues and I all see some Chrimson activation also in non-retinal fed controls, presumably because the intrinsic levels of retinal in the fly are sufficient to drive Chrimson to some degree. Looking through my data, the non-retinal controls are sometimes responding up to about 50% of those fed retinal (this is GCaMP6f response of downstream neurons).
Secondly, flies do see the red light (I use 630nm). It is sometimes suggested that flies can't see far-red light, presumably because the rhodopsin sensitivity doesn't seem to reach much further than 600nm (see Salcedo et al., 1999, JNeurosci), but my colleagues and I have indeed found subtle behavioural as well as neural responses to light >600nm. Presumably Rh6 still has sufficient sensitivity at 630nm if exposed to light with high enough intensity.
So, to sum up, your effects could come from residual Chrimson activity in your controls, or from the light itself. Have you already tested the genetic controls not expressing Chrimson (or any other red-shifted channelrhodopsin, for that matter)?
Cheers,
Oliver
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Hello, recently I've been trying to infect rat lateral amygdala with my AAV5 viral vector, which contains channelrhodopsin or halorhodopsin. 
However, in histological verification, I found repeatedly that it seems like my AAV5 spreads out of its target! Please see the circle area in the attachment. 
It seems like the virus has climbed up along the fiber, but I heard that AAV5 usually does not move in the retrograde direction.
repeatedly, more than 4rats, I found the same expression pattern and I couldn't find why. Is there anybody who experienced the same situation? Or any guess for this?
I'll wait for your comments! Thanks!
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Hello Xinxin, what syringe - cannula system do you use to get an injection volume below 0.5 microL? We use a CR700 Hamilton syringe and connection tubes filled with paraffin oil so that we can use use small tip glass pipettes; but it would be nice to achieve and to control a injection volume like with a nanoliter injector. T
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We are looking at various approaches to inhibit cortico-cortical feedback projections only in mice. Ideally the approach would be source-area specific and target-area specific (only those that project from one area to the other). 
The most straightforward approach would be to inhibit axonal terminals in the target area. However, this is notoriously hard and recently elaborately stated (Mahn et al. 2016 Nat Neurosci). 
Therefore one would like to inhibit the somatas of projecting neurons (e.g. with a retrograde virus without inhibiting other back-projecting regions)
Anyone ideas and/or experience with current approaches?
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rAAV2-retro-Syn-Cre into the target area, then inject flex-[favourite inhibitory opsin] into your secondary area seems like a good place to start. So long as your secondary area is a good few millimeters away, you shouldn't get direct diffusion of the rAAV2-retro, and instead would rely on retrograde transport to bring it to where your flex-inhibitory opsin is.
I use [favourite inhibitory opsin] as opposed to recommending a specific opsin, as I've found both eNpHR3.0 and eArchT3.0 a bit unreliable at shutting down neurons completely. 
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Which laser is most suitable for optogenetic activation of ChR2 in non-neuronal mammalian cells. What power range is required?
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Hi,
Please read this article it sounds they covered different protocols for  activation of channelrhodopsin, however its in nervous cells, but you may optimise the protocol yourself to suit your experiment conditions  
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Hello Everybody, 
We would like to buy wireless optogenetic stimulator for mouse and rat. We found the FireFly and TeleOpto systems, but we don't want to spend lot of time and money unnecessary to gain experience, if someone could share his/her knowledge. If you are using another system contentedly, please let me know.
Thank you for your help, Kornél
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Hi,
Unfortunately nobody answered, I thought you can help me. 
Furthermore Kendall is not very helpful. I asked them, but they did not reply. Today I will Skype with another company, Cambridge Neurotech. In next three month I have to buy some reliable, radio wave guided system, or I lost my grant money.
If you  have any information or suggestion, please share with me.
Bests, Kornél
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I am interested to get ChR2 mammalian expression vectors. Please tell what are the sources I can get hese from. One I know is Addgene. Kindly inform if there are any other too...
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I am trying to develop a rat model for optogenetic stimulation of the basolateral amygdala complex.  Twice now, I have infused a viral vector carrying a channelrhodopsin variant under a CaMKIIa promoter and not seen any expression.  Other rats infused with a control vector (only carrying the fluorophore) do show expression after the same time period post-infusion (i.e. 2 weeks).
The way I see it, there are two options:
1) the viral vector with channelrhodopsin is not functional / dead
2) I have missed the BLA and infused into the lateral ventricle
The latter possibility raises my question: when a viral vector is infused into a ventricle, do you see transfection of the bordering neurons?
I'd appreciate any experience or expertise on the matter.
Thank you!
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Thank you everyone for your advice.
As an update, I just got the histology on another animal in which I injected three distinct vectors in three regions, one being my vector that did not show expression previously.  I injected the problem vector into the sensory cortex this time, and there was still no expression.
I did get expression in the BLA with a different vector (same serotype, same promoter).
Therefore, I believe it is the vector itself that was ineffective, even though the reported titer was high (6 x 10^12 vg/mL).
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If you are doing an experiment where you inject AAV into a specific region, e.g. AAV-DIO-ChR2-eYFP in a Cre driver line for optogenetics, will the AAV transduce the axons passing through the region in addition to the cell bodies? This would compromise a conclusion saying, "Region X neuronal activity is sufficient for Process Y", because really you are activating both Region X and axons from unknown regions. 
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despite initial reports that AAV can be transported in a retrograde fashion, it does so very inefficiently. That's why no one is really using it for this purpose anymore. If you are worried that AAV might be transducing axons, then examine if your transgene is expressed in any cell bodies of these axons - if not than it's not an issue for you. Because for the axon transduction to even matter, the viral particle would need to get it's genome to the nucleus to express it's transgene cargo. I suspect you would find very few if any cell bodies expressing your transgene as a result of axonal viral transduction.
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I read that the serotype 2 tend to spread less than the others. 
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Hi Miguel,
Assuming you are talking about AAVs, we have very good results regarding viral spread and expression in cortical neurons with AAV9. AAV5 also works very well. Even though there are publications showing infection and expression with AAV2 we never had god success with that serotype so I would stay away from it.
Hope this helps,
Ezequiel
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Sometimes my animals will stay put on one corner of the chamber.  Is it ok to repeat a trial with the same animal, while counterbalancing the sides to avoid familiarity with the side-paired-with-stimulus?  I want to give some of the more anxious animals another shot at the task, hoping they will explore the other side of the chamber...
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Hi Alessandra, Thank you for the information!  The difference though is that my experiments are un-conditioned real time place preference...It is a novel exposure experiment with one side paired with optogenetic light activation.  At any rate, these are good resources for me to know in the future if I plan a CPP test! Cheers,
Leandra
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Mostly,  in vivo single-unit recording device with optogenetic activation or inhibtion seems to only use in anesthesia condition. 
Are there some commercial goods for free-moving mouse recording electrophysiology with optogenetics?
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Hi Tsai, I hope you realise this video you linked is a master thesis but not for science but science communication... 
Neuronexus has commercially available optrodes which can be used chronically in freely moving animals. 
In addition, the opeOptogenetics community has these approaches collated:
I hope this helps,
Ede
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I've made optogenetic fiber implants for lasers using lower numerical aperture 0.22 NA silica core fiber 200um from Thorlabs. I used them in Cre-mice injected with AAV-DIO-ChrR2. Now I have to inject mice with ArchT virus. And I wonder if I can use the same optical fibers with 0.22 NA. In the literature I found only people using 0.48 or 0.37NA.
Does anyone know what is the difference between them and what is the impact on the experiment? And if I can use optical fibers of 0.22NA for ARch experiments.
Thanks
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there should be no problem using the same fiber.
higher NA is not needed
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 I injected AAV-ChR2 in rostralPAG and OT-Venus in oxytocin neuron in order to define synaptic connection between them, but none of oxytocin neuron showed light-evoked postsynaptic current.According to previous virus tracing result, they have synaptic connetions. ChR2 expression in axon terminals of PAG neurons seems mild surrounding oxytocin neuron, is that why I couldn't record any photo-responding oxytocin neuron? It wiil be very thankful if you can answer my question!
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In injecting the oxytocin neurons, where did you inject?  Not all OT neurons are similar in terms of the connectivity.
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We are stimulating a part of the cortex with CaMKIIa-ChR2 and in most of the animals, we observe a severe seizure (at least that is what we think it is) that develops after 5 to 10 seconds, including rearing, head shaking, hand-to-mouth movements and sometimes even falling on the side. Even with very low laser stimulation power, we elicited these seizures. Has anyone also already observed this phenomenon and if so, what did you do to prevent it?
Thanks in advance!
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If you pull your fiber too far from your target, you may get false negatives in your behavioral outcome...
Alberto makes an important point.  The seizures could be less a function of your level of stimulation and more related to the brain region where ChR2 is expressed. If you have intentionally or unintentionally included the endopiriform (subregion of the piriform cortex and closely connected to entorhinal) into your list of infected regions, then you could certainly stimulate seizures, as it is a source for seizure activity (see: http://jn.physiology.org/content/86/5/2445.long).  Since you are seeing this even at lower power ranges it starts to rule out activation intensity as a cause.  20mW is a lot of power indeed, but other targeted regions have been published to require 20-50mW as a minimum level of stimulation needed to attain their behavioral effects.  
The endopiriform (in mouse/rat) lies just lateral to the external capsule (see attachment), and that capsule (,"ec" purple arrowhead) acts like a liquid channel, which will conduct your viral injection away from your desired target right down to the endopiriform (black arrow) and possibly the lateral amygdala.  So there could have been unintentional infection of this epicenter of epileptiform activity.  Even if you were mircoinjecting distant structures like the dorsal hippocampus, you can still wind up with virus in the endopiriform with the slightest excess or spillover (by the way, in case you aren't doing this already, be sure to wait 5min after injection is complete before raising your needle, and even then only raise it another 0.2-0.3mm and leave it again for an additional 5min before completely removing your needle to create an air pocket and allow absorption of viral suspension and avoid spillover that can occur with premature removal of your needle).  It would be best to check reporter histology in this area to first confirm/deny this.  
This potential cause I described is all less likely of course, if your target region is much further away (>1mm) as the others have described here.
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Hey, I have a pulser from Prizmatix but i am not sure how to program it to generate different pulses of 1-200 Hz.
it would be nice if someone can have good idea about it.
thanks
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the pulses are defined in miliseconds, rather than HZ.
in order to program a 50 hz pulse, for instance, you need program a 20ms interval and a 20ms pulse duration.
the answer by refik is pretty accurate.
if you need pulses with decimal point timing (2.5ms, for instance) please contact PRIZMATIX support and we will provide you with a software update.
arie
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I'm planning to set up the optogenetic stymulation system in behaving animals and just now I'm considering some software options. I'm currently hesitating between ANY-maze and Ethovision. 
Anybody is using any of these softwares?
I know someone who worked with Ethovision and claims the software does its job correctly but this one is extremely expensive and every other 'module' or upgrade costs few thousands $. ANY-maze seems to be doing similar job for around 7k$ with everything included, but I don't know anybody using it so I don't know if there are some problems that come up only later.
I would very much appreciate any comments concerning your experiences and/or difficulties encountered with both or any of the softwares. 
Thanks!
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A little late to the party, but we use AnyMaze for Opto+Maze tracking simultaneously. If you still have questions let me know and I will do my best to answer. The system works well and the optic fibers rarely cause any issues with tracking.
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I was trying to find an high-power LED to drive the ArchT optogenetically. Two thorlabs LED (590nm and 595nm) have been too weak for us and we found a high power one on prizmatrix. The representative said that this model might cause some noise in Ephys recording. Does anybody have experience with that?
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Hello Shan,
I am using the UHP-Mic-LED-460 from the same company to stimulate my sample during electrophysiological recordings. At first I had some noise from the source, but then when I isolated all the wires that necessarily go into the faraday cage (simply by covering them with aluminium foil and then grounded) the noise disappeared completely. So, in my experience it is not particularly noisy,
good luck
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Hi all. We have been performing in vivo optogenetics for some time now, using DPSS lasers to provide our light source. We find the power of these lasers depreciates quite rapidly (a 100 mW laser can be putting out only 50 mW a few months after purchase), so I would like to know if anyone can recommend a manufacturer that provides good quality, stable lasers. Thanks!
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Hi, the linked ones are reliable manufacturers, but there must be many other. And these lasers can be suitable for you.
If I'm right you measure the laser power after the fiber. Is there any adjustment possibility at the fiber coupling? Its possible that your laser is stil perfect just the coupling needs to be fine tuned.
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Recently, I discovered that there were some abnormal activities of V1 neurons in mice when archaerhodopsin(ArchT) was seletively expressed in parvalbumin(PV)-expressing interneurons.
Spontaneous local field potential of V1 in PV-Cre mice expressing ArchT were different from both in widetype C57BL/6J mice and  SOM-Cre mice expressing ArchT(see the picture attached). This kind of abnormal activity seems not effect on basic response of V1 neurons to visual stimuli, that's to say, receptive field and orientation tuning could be getted.
 But, after the V1 cortex was exposed in green laser for sometime, spontaneous local field potential of V1 without laser will exhibit another kind of abnormal activity(also see attached picture). That seems that laser effect V1 neurons irreversibly,  inconsistent with the reversible effect of optogenetics. And visual stimuli can aggravate this kind of abnormal activity, which  seriously disturbing response of V1 neuron to visual stimuli, even that you can not get the receptive of V1 neurons. It's less happened in SOM-Cre mice expressing ArchT
Is it happened in your experiments? Do you know why this happen? And how to improve the situation?
Look forward to your reply
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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).
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Our lab is in the process of setting up in vivo optogenetics in RATS for behavioral studies. The goal is to activate specific projections based on their connectivity, so we are considering the method of injecting the opsin into region A and a WGA-CRE virus into region B, so that when we stimulate A, only those cells projecting to B get activated. 
Is this method reliable enough to get good expression of the opsin? Are there issues with spillover? We are planning on using AAV5 for the opsin to get a large region of transfection. Is AAV5 the best serotype to use for WGA-CRE as well, or would something like AAV9 work better?
How does this method compare with using CRE packaged in a retrograde virus (e.g., CAV, PRV)?
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It sounds like the CAV-Cre + AAV-DIO-opsin virus combo would be best.
I've used WGA alone as a tracer in the brain and via virus as AAV-WGA-Cre, and it kind of goes everywhere. WGA is quickly transported transsynaptically retrogradely and anterogradely through most neural circuits. Thus, the issue that occurs is whether the DIO construct is being expressed in first, second, or even third order neurons in relation to the AAV-WGA-Cre injection site. For this reason I'd avoid AAV-WGA-Cre, regardless of serotype.
Replication-incompetent PRV would work better as it would go monosynaptically retrogradely (depending on serotype, I think), but it requires biosafety level 2 facilities and quarantine protocols. I also haven't heard of labs that distribute PRV-Cre viruses, though they may exist.
The CAV virus infects cells monosynaptically retrogradely, it only requires BSL1 facilities, and there is a distributor for it in France. I personally haven't used it but I know people at my university that have used it in combination with a Cre-dependent AAV construct (DIO-DREADD instead of opsin) with reasonable success. The timeline is the same as a non-Cre dependent AAV for optimal opsin expression, about 4-6 weeks post-injection.
Good luck!
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I would like to optogenetically silence specific neurons by retrogradely labeling them using a viral vector. Im thinking of using an AAV-Serotype 6 virus with ArchT. Anybody knows if such a vector exists or have experience with it? Any help/advice would be appreciated
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CAV-2 is good at retrograde labeling :  http://www.igmm.cnrs.fr/spip.php?rubrique166
If you still want to use AAV 6, our virus core can produce some for you (but it's not a stock virus):
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Leaning toward serotype 5. I have read that serotype 5 is better in adult rodents. That was outlined here
Han X. Optogenetics in the nonhuman primate. Progress in brain research. 2012;196:215-233. doi:10.1016/B978-0-444-59426-6.00011-2.
Seeing if anyone has experience handling these vectors and injecting them.
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Ok, just so you know, I've used about every serotype under the sun, in the VTA and elsewhere. Here's the deal in a nutshell:
AAV9: has anterograde and retrograde properties. One of the early experiments with it tried to used the VTA as a sort of highway to achieve the highest degree of transduction in the brain (i.e. most brain regions, widest spread, etc) for treatment of disorders that affect neurons globally. You do NOT want to use this, unless you know that none of the neurons projecting to the VTA contain the promoter driving Cre (TH wouldn't be good, since there are some noradrenergic projections to the VTA).
AAV5: can transduce both neurons and glia (as can many others including AAV2 and AAV8), but it's preferentially neuronal, and many of the papers pushing glial expression used a glial specific promoter, so the neuronal transduction didn't show. My understanding is Deisseroth prefers AAV5 because of the issue with the ChR2 blebbing. Blebbing isn't good for the neurons, AAV5 has higher expression but not TOO high, so the blebbing is minimal while still driving enough expression to be behavioral or physiologically relevant.
AAV2: expression is so poor it's not worth trying. I would recommend anything over this, even at massive volumes you won't get good expression, mostly the needle track. You wind up super infecting a small populations of neurons, it just doesn't spread. If you want more spread though you can co-infuse with heparin, but again, it's generally not worth your time. 
AAV8: also very good, similar to 5, may be a bit stronger. 
AAV10: my preferred serotype for the brain, easily titratable, I've restricted to the SCN of a mouse with 50 nl and gotten nearly half with the striatum with 0.5 ul. I've injected in mice, rats, prairie voles, primates, it gives really nice expression across the board. And of course I've published 3-4 papers with it in the VTA to express ChR2. But I've published at least a dozen in the CNS in general.
I make all of the AAVs in my lab. I would be very cautious when dealing with core facilities, for reasons I'm not willing to go into here. Feel free to contact me if you want some advice. Cores are nice just keep your ears open to any problems. 
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Some paper said that,'Clumping of a channelrhodopsin
fused to mCherry  was apparent , suggesting that mCherry may not be an ideal
fluorophore for opsin usage'.So I'm  considering if it is necessary to reconstruct the carrier,pAAV-EF1a-double floxed-hChR2(H134R)-mCherry into pAAV-EF1a-double floxed-hChR2(H134R)-EGFP. What's yoursuggestion?
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 Hi, Dr. Volker Busskamp,
Thanks for your reply and suggestion. We are going to buy  some virus carrying pAAV-EF1a-double floxed-hChR2(H134R)-eYFP to apply. And  I think it is a wise decision to test pAAV-EF1a-double floxed-hChR2(H134R)-mCherry by myself in my experiment as you said. So I will also try it.Thank you again.