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.
Questions related to Optogenetics
Why does the laser for optogenetic stimulation need to be shuttered on and off rapidly? Can i use a constant laser light but at low power?
I am in the doing viral injection (hamilton and automatic pump) with AAV-mcherry targeting piramidal neurons in mouse aging from P40 to P50. After 2 to 3 weeks, I performed optogenetics stimulation, then after having the brain removed, soaked in formalin and fixated for at least 48h, I checked the viral expression with fluorescence microscope.
Here's the point: out of 10 trials, 4 times I didn't find any trace about the viral expression. I have checked the whole brain, but still zero. We are talking about zero level expression. On the other hand, in 6 experiements I have strong and proper expression.
In my experimental routine, I had 2 injections per day (2 mice). On the same days, I withdraw the virus out of the same small eppenderf tube each time for one animal. Between the two viral injections, the virus is kept soaked in ice in a dark water proof container.
I am sure the virus was injected (I saw the virus level in the glass capillary lowering). As always, I have waited 10 min before needle removal.
What could have happened in those 4 trials with no expression? Is the virus contained in the eppendorf damaged (at least in some of them)? I really don't know what to think right now.
Any help would be much appreciated.
I am just curious if there will be changes in function/behaviour if you optogenetically activate a particular region of the neuron (e.g. dendrite), compared to activating the whole neuron itself?
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!
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?
I have started working in the field of optogenetics and I have photonics background. I would like to understand the basis on which an opsin is selected for optogenetic experiments. Moreover, I would also like to learn about choosing the right DNA construct with fluorophores and promoter (including their function etc).
I could only find scattered data online. Can someone help me with any literature or textbooks that explain these concepts with examples for someone who doesn't have biology background?
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
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!
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.
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.
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.
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)?
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!
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...
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.
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.
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.
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.
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
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.
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.
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!!
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.
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.
If a neuron expressing an optogenetic actuator is light-stimulated in the soma, will this lead to neurotransmitter release of this same neuron?
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
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.
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)
Please provide solution that is readily available. Also what is the downsides of each of them when comparing to electrophysiology?
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?
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.
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.
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.
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 !
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[
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!
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 :)
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.
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.
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!
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.
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.
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.
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.
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.
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!
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
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.
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?
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?
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.
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?
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
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...
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.
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.
I read that the serotype 2 tend to spread less than the others.
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...
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?
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.
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!
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!
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.
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.
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?
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!
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
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)?
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